Plant Soil (2009) 321:5–33
DOI 10.1007/s11104-009-9925-0
REVIEW ARTICLE
Carbon flow in the rhizosphere: carbon trading
at the soil–root interface
D. L. Jones & C. Nguyen & R. D. Finlay
Received: 30 July 2008 / Accepted: 4 February 2009 / Published online: 25 February 2009
# Springer Science + Business Media B.V. 2009
Abstract The loss of organic and inorganic carbon
from roots into soil underpins nearly all the major
changes that occur in the rhizosphere. In this review
we explore the mechanistic basis of organic carbon
and nitrogen flow in the rhizosphere. It is clear that C
and N flow in the rhizosphere is extremely complex,
being highly plant and environment dependent and
varying both spatially and temporally along the root.
Consequently, the amount and type of rhizodeposits
(e.g. exudates, border cells, mucilage) remains highly
context specific. This has severely limited our
capacity to quantify and model the amount of
rhizodeposition in ecosystem processes such as C
sequestration and nutrient acquisition. It is now
evident that C and N flow at the soil–root interface
is bidirectional with C and N being lost from roots
Responsible Editor: Philippe Hinsinger.
D. L. Jones (*)
School of the Environment & Natural Resources,
Bangor University,
Bangor, Gwynedd LL57 2UW, UK
e-mail: [email protected]
C. Nguyen
INRA, UMR1220 TCEM,
71 Avenue Edouard Bourlaux, BP 81,
33883 Villenave d’Ornon, France
R. D. Finlay
Uppsala BioCenter,
Department of Forest Mycology and Pathology, SLU,
Box 7026, SE-750 07, Uppsala, Sweden
and taken up from the soil simultaneously. Here we
present four alternative hypotheses to explain why
high and low molecular weight organic compounds
are actively cycled in the rhizosphere. These include:
(1) indirect, fortuitous root exudate recapture as part
of the root’s C and N distribution network, (2) direct
re-uptake to enhance the plant’s C efficiency and to
reduce rhizosphere microbial growth and pathogen
attack, (3) direct uptake to recapture organic nutrients
released from soil organic matter, and (4) for interroot and root–microbial signal exchange. Due to severe
flaws in the interpretation of commonly used isotopic
labelling techniques, there is still great uncertainty
surrounding the importance of these individual fluxes
in the rhizosphere. Due to the importance of rhizodeposition in regulating ecosystem functioning, it is
critical that future research focuses on resolving the
quantitative importance of the different C and N fluxes
operating in the rhizosphere and the ways in which
these vary spatially and temporally.
Keywords Carbon cycling . Nitrogen cycling .
Mycorrhizas . Organic matter . Review .
Rhizodeposition . Root processes . Signal transduction
Introduction
For over a century it has been established that plants
can dramatically modify their soil environment giving
rise to the so called rhizosphere effect (Clark 1949;
6
Rovira 1965; Whipps 2001). Although the initial
trigger of this rhizosphere effect was not identified,
subsequent research has shown that it is largely
induced by the release of carbon (C) from roots into
the surrounding soil. Although roots can release large
amounts of inorganic C which may directly affect the
biogeochemistry of the soil (Cheng et al. 1993;
Hinsinger 2001; Hinsinger et al. 2009), it is the
release of organic carbon that produces the most
dramatic changes in the physical, biological and
chemical nature of the soil. In its broadest sense, this
release of organic C is often termed rhizodeposition
(Jones et al. 2004). The term rhizodeposition includes
a wide range of processes by which C enters the soil
including: (1) root cap and border cell loss, (2) death
and lysis of root cells (cortex, root hairs etc), (3) flow
of C to root-associated symbionts living in the soil
(e.g. mycorrhizas), (4) gaseous losses, (5) leakage of
solutes from living cells (root exudates), and (5)
insoluble polymer secretion from living cells (mucilage; Fig. 1). Although these loss pathways can be
clearly differentiated between at a conceptual level it
is often extremely difficult at the experimental level to
discriminate between them in both space and time.
Consequently, while individual studies have shown
that these can all occur, probably simultaneously in
the same plant root system, it is almost impossible to
rank the relative importance of each process. Further,
as we understand more about the mechanisms of C
flow in both soil and roots we find that many of the
published results are severely biased by the experimental system in which individual factors or processes were examined (Jones and Darrah 1993; Meharg
1994; Kuzyakov 2006). This has left the literature on
rhizosphere C flow awash with studies which may
bear no relationship to real world events, particularly
those performed in the absence of soil. Despite this,
however, it is clearly apparent that our incremental
approach to understanding C flow is paying dividends
from both a commercial and environmental perspective. Firstly, from a commercial perspective it is clear
that root C excretions can be useful for the nondestructive production of high value pharmaceuticals,
pigments and flavours for use in the medical and
cosmetic industries (Oksman-Caldentey and Inze
2004). In these applications, roots are typically transformed with Agrobacterium rhizogenes which induces hairy root disease. The neoplastic (cancerous)
transformed roots are genetically stable and can grow
Plant Soil (2009) 321:5–33
6
5
4
3
1
2
Fig. 1 Schematic representation of a longitudinal section of a
growing root showing the six major sites of rhizodeposition: 1
loss of root cap and border cells, 2 loss of insoluble mucilage, 3
loss of soluble root exudates, 4 loss of volatile organic C, 5 loss
of C to symbionts (e.g. arbuscular mycorrhizas), and 6 loss of C
due to death and lysis of root epidermal and cortical cells
rapidly in the absence of shoots in a hormone free
medium making them suitable for the controlled
excretion and collection of secondary metabolites
(Srivastava and Srivastava 2007). In our efforts to
create a more sustainable environment, it is also clear
that rates of release of C compounds from roots can
be manipulated to increase food production, enhance
water conservation, speed up the remediation of
contaminated sites, and reduce the need for artificial
fertilizers and pesticides (Lasat 2002; Vessey 2003;
Welbaum et al. 2007). Thus while the intricate
complexity of the rhizosphere continues to amaze us
and presents a real challenge to scientists trying to
unravel its diverse web of interactions, it also has the
potential to offer great benefits to society. As C flow
from roots is essentially the starting point from which
the rhizosphere develops it is important that we
improve our understanding of this process.
Plant Soil (2009) 321:5–33
7
This review aims to critically assess our current
understanding of rhizosphere C flow and to highlight
areas for further research. Due to the large number of
publications in this research area (>5000 individual
publications) it is not our aim to cover the entire literature
but to highlight examples to support themes. Readers
requiring more comprehensive historical reviews of the
literature should consult Rovira (1969), Curl and
Truelove (1986), Lynch (1990) and Pinton et al. (2001).
Roots release a great variety of compounds
by different mechanisms
Virtually, all compounds contained in root tissues can be
released into the soil. In hydroponic culture, carbohydrates, organic and amino acids, phenolics, fatty acids,
sterols, enzymes, vitamins, hormones, nucleosides have
been found in the root bathing solution (Rovira 1969;
Grayston et al. 1996; Dakora and Phillips 2002; Read
et al. 2003; Leinweber et al. 2008). These compounds
are released by various mechanisms including secretion, diffusion or cell lysis. Depending upon the
question being addressed, different nomenclatures for
rhizodeposits have been proposed based for instance
on the mechanisms of release, on the biochemical
nature of rhizodeposits or on their functions in the
rhizosphere. The classification first proposed by Rovira
et al. (1979) is generic and has been extensively used.
Mucilage
Root mucilage forms a gelatinous layer surrounding
the root tip and is one of the few clearly visible signs
of organic C excretion from roots (Fig. 2). It is mainly
composed of polysaccharides of 106–108 Da in size
(Paull et al. 1975) and is actively secreted by
exocytosis from root cap cells (Morre et al. 1967;
Paull and Jones 1975a, b, 1976a, b). Alongside
polysaccharides, it also contains proteins (ca. 6% of
dry weight; Bacic et al. 1987) and some phospholipids (Read et al. 2003). In most situations mucilage
released into the soil confers a wide range of benefits
to the plant. For example, the carboxylic groups of
mucilage can complex potentially toxic metals (e.g.
Al, Cd, Zn, Cu), protecting the root meristem (Morel
et al. 1986; Mench et al. 1987). In addition, mucilage
enhances soil aggregate stability which in the longterm promotes soil aeration, root growth and reduces
Fig. 2 Light microscope image showing the large amount of
mucilage (blue halo surrounding the root) and border cells
production in a Zea mays L. root tip. Labels indicate the root
quiescent centre (A), the main root elongation zone (B), and the
mucilage halo in which the border cell are embedded (C). The
mucilage is stained with aniline blue
soil erosion (Guckert et al. 1975; Morel et al. 1990;
Czarnes et al. 2000). Mucilage also possesses a high
intrinsic affinity for water and when fully hydrated,
has a water content 100,000 times greater than its dry
weight (McCully and Boyer 1997) and expands to
form a viscous droplet covering the root tip. Such
properties play a role in maintaining the continuity of
water flow towards the rhizoplane (Read and Gregory
1997; Read et al. 2003) and in reducing the frictional
resistance as the root tip moves through the soil
(Iijima et al. 2004). Recent work has also suggested
that specific mucilage components (e.g. prenylated
stilbenes) possess antimicrobial properties and may be
important in preventing pathogen attack (Sobolev et
al. 2006). Apart from the C employed to synthesize
and secrete mucilage, its loss into the soil appears to
have no known negative effects on soil and plant
health. Of most concern is that mucilage represents a
source of labile C in the soil and is consequently
rapidly consumed by soil microorganisms (typical
half-life of 3 days). In some instances this can induce
the proliferation of root rot disease-causing organisms
in the rhizosphere (e.g. Pythium aphanidermatum;
Zheng et al. 2000) while in other situations due to its
high C:N ratio (ca. 65), its biodegradation induces a
transient net immobilisation of N in the rhizosphere
(Mary et al. 1993; Nguyen et al. 2008). While
8
increasing the rate of polysaccharide mucilage release
from roots is possible and would only constitute a
minor C drain (Darrah 1991a), it is unlikely to yield
major benefits in comparison to alteration in other
rhizodeposition processes.
Border cells
Also called sloughed-off cells, border cells are the
cells that detach from the external layers of the root
cap, which is continuously renewed (Barlow 1975;
Fig. 2). The controlled separation of the border cells
reduces the frictional force experienced by the root tip
(Bengough and McKenzie 1997; Bengough and
Kirby 1999; Iijima et al. 2004). The daily rate of
border cell production is highly variable among plant
species, from none to tens of thousands with release
rates highly dependent upon the prevailing environmental conditions (Hawes et al. 1998; Iijima et al.
2000; Zhao et al. 2000). Once detached from the cap,
border cells remain alive in the soil for several days
(Stubbs et al. 2004). They are surrounded by the
mucilage they secrete, which binds heavy metals away
from the root meristem (Miyasaka and Hawes 2001).
Border cells also produce signal compounds involved
in the protection of meristem against pathogens
(Hawes et al. 2000) and in the promotion of symbiosis
(Brigham et al. 1995; Hawes et al. 1998). Recent work
has also suggested that border cells can act as a decoy
luring pathogenic nematodes and fungi away from the
main root axis (Gunawardena and Hawes 2002;
Rodger et al. 2003). However, contradictory results
have also been found highlighting the difficulties of
manipulating border cell release and physiology for
disease control (Wuyts et al. 2006; Knox et al. 2007).
While border cells may provide a convenient mechanism for compound delivery to soil, further fundamental work is required to characterise the metabolomic
and proteomic expression patterns in comparison to
other root cells to understand and capitalize on their
unique attributes (Jiang et al. 2006). In the total
rhizodeposition C budget, however, border cells only
constitute a small proportion of the C entering the soil
(Iijima et al. 2000; Farrar et al. 2003).
Exudates
Exudates are defined as diffusible compounds which
are lost passively by the root and over which the root
Plant Soil (2009) 321:5–33
exerts little direct control. Rates of loss of individual
compounds depend upon three critical factors, namely
(1) the root-soil concentration gradient, (2) the
permeability of the plasma membrane, and (3) the
spatial location of the solutes in the root tissue (e.g.
epidermis versus stele). The dominant organic compounds in roots reflect those compounds central to
cell metabolism and include free sugars (e.g. glucose,
sucrose), amino acids (e.g. glycine, glutamate) and
organic acids (e.g. citrate, malate, oxalate; Kraffczyk
et al. 1984). Their concentration inside the root is
typically orders of magnitude greater than that in the
surrounding soil solution due to continual removal
from the soil by the soil microbial community and
replenishment of internal pools by the root. Although
we know a great deal about the concentrations of
solutes in whole roots our understanding of the spatial
and temporal dynamics of organic solutes in roots is
severely limited. As there is significant internal
partitioning of solutes in root cells (e.g. cytoplasm
versus vacuole; Gout et al. 1993; Ciereszko et al.
1999), of critical importance is the actual concentration gradient that exists between the cytoplasm and
the cell wall space rather than that between the whole
root and bulk soil. In addition, at a tissue level, the
role of the cortex in root exudation versus that of the
epidermis remains unknown. Although apoplastic loss
may represent a slow diffusion pathway in comparison to direct loss from the epidermis (Canny 1995;
Fleischer and Ehwald 1995), evidence suggests that
gaps between epidermal cells (where apoplastic loss
ultimately manifests itself) are strong regions of
microbial colonization and therefore C availability
(Quadt-Hallmann et al. 1997; Watt et al. 2006).
Therefore more work is required to characterise
apoplastic loss pathways from the root cortex and its
contribution to maintaining the endo- and ectorhizosphere microbial community. Further work is
also required to determine the temporal dynamics of
solutes in root tissues (e.g. diurnal versus ontogenetic)
and the relationship with exudation.
The cytoplasmic pH of most root cells ranges from
7.2–7.5. Within this range most organic acids are
negatively charged while most amino acids and
sugars carry no net charge. Due to plasma membrane
H+-ATPases pumping H+ out of the cells, the outside
of the plasma membrane carries more positive charge
than the inside (Fig. 3). Consequently, there is a
greater tendency for anionic organic solutes to be
Plant Soil (2009) 321:5–33
9
= glucose
Proton pump
(H+-ATPase)
H+
Outside
the root
Inside
the root
H+-sugar
co-transporter
H+
1
ATP
3
1
ADP + P i
H+
H+
Fig. 3 Schematic representation of the three main processes
involved in the bi-directional flux of low molecular weight
organic solutes (e.g. glucose) across the soil root interface. Flux
(1) denotes the passive transport of glucose across the plasma
membrane in response to the large cytoplasm (20 mM) to soil
solution (10 μM) concentration gradient. Flux (2) denotes the
active energization of the plasma membrane by the H+-ATPase
which pumps H+ out of the cell using ATP as the energy source.
Flux (3) denotes the active re-uptake of sugars from the soil
solution back into the cytoplasm using a H+-cotransport
protein. The cell wall is not drawn for clarity
drawn across the membrane at faster rates than noncharged solutes (Ryan et al. 2001). Studies in nonplant systems suggest that although solutes can
diffuse through the lipid bilayer, faster rates of
diffusion occur at the lipid–protein boundary. Further,
organic solute loss may be accelerated at sites where
active growth is occurring as membrane vesicle
contents are released during fusion with the plasma
membrane. Rates of exudation can also be greatly
speeded up by the opening of solute specific channels
in the membrane. Probably the best known example
of this is the release of organic acids when roots
experience either P deficiency or high external
concentrations of free toxic Al3+ (Zhang et al. 2004;
Ligaba et al. 2006). The release of organic acids such
as citrate, malate and oxalate can complex the Al3+
rendering it non-toxic. Detailed reviews of the role of
organic acid channels in metal detoxification and
nutrient uptake (e.g. P) can be found in Ryan et al.
(2001), Jones et al. (2004) and Roberts (2006).
Generally, rates of exudate loss are greater at root
tips in comparison to mature root regions (McDougal
and Rovira 1970; Hoffland et al. 1989). Potential
reasons to explain this enhanced C loss from tips
include: (1) higher solute concentrations in root tip
regions thereby creating a larger diffusion gradient
(Jones and Darrah 1996; Jones et al. 1996), (2) small
vacuolar volume of root tip cells inducing higher
cytoplasmic concentrations (Patel et al. 1990), greater
surface area-to volume ratio of tip cells, (3) the lack
of an endodermal layer to minimize cortical loss
(Schraut et al. 2004), (4) increased rates of apoplastic
solute unloading from the vascular tissue leading to
greater apoplastic loss (Bockenhoff et al. 1996), (5)
greater apoplastic volume inducing higher rates of
solute diffusion (Kramer et al. 2007), (6) higher rates
of growth in tip regions and therefore solute loss
during vesicle fusion and signalling events (Beemster
and Baskin 1998; Roux and Steinebrunner 2007), and
(7) localized loss of root border and cap cells which
may undergo apoptosis releasing solutes (Shishkova
and Dubrovsky 2005). Like many other aspects of
rhizodeposition our conceptual understanding is good,
however, our detailed mechanistic understanding of
10
the relative importance of the individual flux pathways remains poor and this must remain as a priority
for future research.
Secretions (excluding mucilage)
Plant roots actively secrete various compounds in
response to a range of environmental conditions and
our understanding of the role of these compounds in
rhizosphere processes often remains poor (Wen et al.
2007). One exception is the characterisation of
phytosiderophore release by grasses under conditions
of low Fe availability (Negishi et al. 2002). In this
situation, Fe-phytosiderophore complexes are also
actively taken back into the plant (Haydon and
Cobbett 2007). Phenolics are also secreted from roots
and have been implicated in the mobilization of
nutrients such as Fe and P, however, their quantitative
importance remains unknown (Dakora and Phillips
2002). Enzymes (e.g. phosphatase) and many other
compounds such as secondary metabolites may also
be secreted into the rhizosphere and participate in the
interactions between the roots and their environment
(Bais et al. 2004). High molecular weight compounds
or toxic molecules are likely to be released by
exocytosis (Verpoorte et al. 2000). However, the
mode of release is not always clearly established
and much further work is required to elucidate
mechanisms of release and their quantitative significance in the soil.
Plant Soil (2009) 321:5–33
the relative importance of these two processes.
However, we do know that plant roots contain a
significant amount of soluble and insoluble C and that
their death will results in a significant C and N input
to the soil and an elevation of microbial populations
in their necrosphere (McClaugherty et al. 1982;
Nadelhoffer and Raich 1992; Stewart and Frank
2008). Typically, there is a positive correlation
between root diameter and lifespan (Gill and Jackson
2000). Consequently, in temperate agricultural grasslands containing an abundance of fine roots, we can
calculate the magnitude of the C input to soil from
root turnover. The soil organic C content of a
temperate, grazed grassland soil typically ranges from
10 to 50 g C kg soil−1 while the standing root biomass
typically ranges from 5 to 15 g root-C kg soil−1 and
the microbial biomass from 0.5 to 1 g C kg soil−1 of
which we assume only 10% is active (Jones,
unpublished). It has been estimated that in the
growing season approximately 25% of the roots turn
over each month equating to approximately 2 to 10 g
C kg soil−1 month−1 (i.e. enough C to generate 50 to
100 times the size of the active microbial biomass in
soil). This can be compared to the rates of C
exudation from grass roots which typically range
from 1 to 10 mg C g root-C−1 day−1 (Hodge et al.
1997; Paterson and Sim 1999; Paterson et al. 2003).
Consequently, we can estimate the amount of C
entering grassland soils from root exudation to be in
the range 0.1 to 5 g C kg soil−1 month−1 similar to
that derived from root turnover.
Senescence-derived compounds
Carbon flow to mycorrhizal and bacterial symbionts
Depending upon the conditions experienced by the
root, a variable part of the epidermis including root
hairs and of the cortical cells can degenerate and
release their content into the rhizosphere (Fusseder
1987; McCully 1999). As roots rarely senesce in
hydroponic culture, this process is largely thought to
occur in soil where pathogens and mineral abrasion
can induce cell death. Little is known about the
magnitude of this flux pathway as it is almost
impossible to study in soil. Consequently, most
measurements typically rely on quantifying the
amount of epidermal and cortical cell loss rather than
the amount of C transferred to the soil. The amount
entering the soil can be expected to depend upon
whether the roots undergo programmed (apoptosis) or
spontaneous cell death, however, little is known about
Apart from the bacterial-legume symbiosis which has
been reviewed extensively, little is known about the
flow of C to other bacterial symbionts in the
rhizosphere (Dilworth et al. 2008; Ohyama et al.
2009). Consequently, here we will focus on mycorrhizas. Most plants in natural and semi-natural
vegetation systems form symbiotic associations with
mycorrhizal fungi and there is increasing evidence to
suggest that the flow of C to and through this
symbiotic interface may be of significance in many
plant–soil interactions, playing an important role in
different biogeochemical processes (Finlay and Rosling
2006; Finlay 2008). Mycorrhizal symbionts contribute to carbon flow in the rhizosphere in three main
ways. Firstly, the investment of C in production of
Plant Soil (2009) 321:5–33
biomass of intra- and extraradical mycelial structures is,
in itself, substantial (Leake et al. 2004). Secondly, there
is a flow of C through these structures, resulting in
release of a range of exudates into the mycorrhizosphere, and thirdly, these compounds, and the mycorrhizal mycelium itself, can be used as energy rich
substrates by other organisms, resulting in respiratory
loss of carbon as CO2. As with studies of other
components of rhizodeposition, considerable effort has
been directed at quantifying the contribution of these
processes in relation to total rhizosphere C flow, whilst
fewer studies have focused on their potential functional
roles.
Because of their fine dimensions and fragility,
mycorrhizal hyphae are even more difficult to study
than fine roots. The mycelium is easily damaged
when excavating roots, it consists of viable and nonviable fractions and must be distinguished from the
mycelia of saprotrophic and pathogenic fungi. Despite
these difficulties, much knowledge has been gained
about the structure, biology and impact of mycorrhizal mycelia (see Leake et al. 2004 for an extensive
review). Over 50 estimates of mycelial production by
arbuscular mycorrhizal (AM) fungi or ectomycorrhizal (EM) fungi are cited from a range of pot and field
studies. Estimates of hyphal length for AM fungi
typically range from 3–30 m g−1 soil but 68–101 m
g−1 soil have been recorded in undisturbed grasslands
with permanent plant cover. EM hyphae are more
difficult to distinguish morphologically from saprotrophic fungi and hyphal length estimates are less
reliable but available data suggest hyphal length
densities of between 3 to 600 m g−1 soil. Wallander
et al. (2001) used a combination of techniques such as
in-growth mesh bags, measurements of fungal markers
such as phospholipid fatty acids and ergosterol, δ13C
values and trenching to distinguish mycorrhizal fungi
from soil dwelling saprotrophs. The total amount of
EM mycelium colonising the mesh bags was calculated
to be 125–200 kg ha−1 and the total amount of EM
mycelium, including EM mantles was estimated to be
700–900 kg ha−1.
Clearly the investment of C in mycelial structures
is considerable and many attempts have been made to
estimate C allocation to mycorrhizal mycelium. Many
of these involve labelling studies with radioactive or
stable isotopes and are subject to different sources of
error. Microcosm studies may result in unnaturally
high mycelial biomass and/or exclude soil biota
11
which may graze fungal mycelia. However, short
pulse-labelling experiments may underestimate C
allocation to mycorrhizal mycelia since they only
measure cytoplasmic allocation and exclude C allocation to previously formed fungal cell walls. Many
experiments fail to measure respiratory losses of
labelled CO2 which complicates the construction of
complete C budgets. C flow through arbuscular
mycorrhizal (AM) mycelia has been measured in
grassland ecosystems dominated by AM mycelia and
found to be at least as large as that of fine roots, with
at least 5.4–7.7% of the C lost by plants being
respired from AM fungal mycelium and 3.9–6.2%
being fixed in mycorrhizal mycelium within 21 h of
labelling (Johnson et al. 2002). These figures are
comparable with those for fine roots and suggest that
there is a very rapid flux of C through mycorrhizal
hyphae. A particular strength of these data is that they
were obtained under field conditions. Additional
studies using similar methods have investigated the
effects of soil invertebrates and shown that they can
disrupt C transport through hyphal networks but that
there is still a significant, rapid flow (Johnson et al.
2005). Analyses of 14C content of AM hyphae by
accelerator mass spectrometry (Staddon et al. 2003)
suggest that most hyphae live for 5–6 days, again
suggesting that there is a large and rapid pathway of C
flow through the AM extraradical mycelium.
Measurements of C flow to ectomycorrhizal
mycelium colonising forest trees are more difficult
to obtain due to the size of the plant hosts, but data
from smaller plants in microcosm systems (Leake et
al. 2001) showed that the extraradical mycelium of
the ectomycorrhizal fungus Suillus bovinus colonising
Pinus sylvestris seedlings contained 9% of the 14C
contained in the plants 56 h after labelling. Over 60%
of the C allocated to the extraradical mycelium was
allocated to mycelium colonising patches of litter,
which only represented 12% of the available area for
colonisation, suggesting that this C allocation was
associated with nutrient acquisition. Data from a
range of microcosm-based labelling studies (see
Leake et al. 2004 for details) suggest that 7–30% of
net C fixation is allocated to ectomycorrhizal mycelium and that 16–71% of this C is lost as respiration.
These data are likely to be underestimates of C
transfer to the mycelium since short term pulselabelling experiments do not measure the carbon in
the fungal cell walls. Although microcosm experi-
12
ments may not accurately reflect field conditions,
manipulation of the ectomycorrhizal extraradical
mycelium in forest ecosystems using the methods
employed by Johnson et al. (2002) is not possible due
to the large size of the plants. Tree girdling experiments in a 45–55 year old pine forest by Högberg et
al. (2001), however, suggest that soil respiration is
directly coupled to the flux of current assimilate to
mycorrhizal roots and fungi. Decreases of 37% were
recorded within 1–2 days, however, this method does
not allow separate determination of the root and
fungal components. Further observations following a
large-scale girdling experiment suggest that ectomycorrhizas may contribute at least 32% of soil
microbial biomass and as much as half the dissolved
organic carbon in forest soil (Högberg and Högberg
2002). The below-ground flux of recent photosynthate
has been followed with high temporal resolution using
13
C labelling of 4-m-tall Pinus sylvestris trees (Högberg
et al. 2008). C in the active pools in needles, soluble
carbohydrates in phloem and in soil respiratory efflux
had half-lives of 22, 17 and 35 h, respectively. C in
soil microbial cytoplasm had a half-life of 280 h,
while the C in ectomycorrhizal root tips turned over
much more slowly. Simultaneous labelling of the soil
with 15NH4+ showed that the ectomycorrhizal roots,
which were the strongest sinks for photosynthate,
were also the largest sinks for N. Tracer levels
peaked after 24 h in the phloem, after 2–4 days in
the soil respiratory efflux and soil microbial cytoplasm and 4–7 days in the ectomycorrhizal roots. The
results indicate close temporal coupling between tree
canopy photosynthate and soil biological activity.
Other recent studies using free air carbon dioxide
enrichment (FACE) experiments as a means of 13C
labelling (Körner et al. 2005) and bomb 14C estimates
of root age (Gaudinski et al. 2001) suggest that fine
roots of trees may turn over much more slowly than
previously assumed. This suggests that more of the
below-ground C flux may take place through mycorrhizal fungi and other soil biota associated with roots
(Högberg and Read 2006). A recent FACE study of a
Populus plantation supports this idea, suggesting that
extraradical mycorrhizal mycelium is the dominant
pathway (62%) through which C enters the soil
organic matter pool (Godbold et al. 2006).
Both arbuscular mycorrhizal and ectomycorrhizal
plants can regulate their C allocation to roots.
Trifolium repens plants have been shown to increase
Plant Soil (2009) 321:5–33
their rates of photosynthesis in response to increased
sink strength of mycorrhizal roots and to increase
activities of cell wall and cytoplasmic invertases and
sucrose synthase (Wright et al. 1998). In ectomycorrhizal plants the symbiotic partners receive up to 19
times more carbohydrates from their roots than
normal leakage would cause, resulting in a strong C
sink. To avoid parasitism the plants appear to have
developed mechanisms to regulate the C drain to the
fungal symbiont in relation to the supply of fungusderived supply of nutrients (Nehls 2008). Increased
expression of plant and fungal hexose transporter
genes has been detected at the plant fungus interface
in ectomycorrhizas, but it appears there may also be
mechanisms to restrict carbohydrate loss to the
fungus. Hexoses generated from sucrose hydrolysis
by plant-derived acid invertases could be taken up by
plant or fungal cells through monosaccharide transporters. One Poplar sugar transporter gene (PttMST3.1)
is expressed at least 10 times more highly than other
hexose transporter genes and it is postulated that this
may be regulated at the post-transcriptional level by
phosphorylation which would allow activation of the
transporter as a reaction to the amount of nutrients
delivered by the fungus. If the fungus provided
sufficient nutrients the activity of the transporter
would be shut off, while the protein would be
activated as soon as the nutrient transfer is insufficient (Nehls 2008). Unpublished data support this
hypothesis but further studies of the genetic basis of
regulation of carbon flow at the symbiotic interface
are still needed in a range of different mycorrhizal
associations.
One disadvantage of simple labelling experiments
showing transport of a labelled element from a source
to a sink is that they provide no information about net
movement of the element in question, since there may
be an equal (or greater) movement of the same
(unlabelled) element in the reverse direction. The
issue of C transport between plants connected by a
common mycorrhizal mycelium has been controversial. Experiments by Francis and Read (1984)
demonstrated the potential for transfer of C along
concentration gradients from sources to sinks induced
by shading, however, these studies were criticised for
the above reasons. Experiments by Simard et al.
(1997) using reciprocal labelling with 14C and 13C
demonstrated net transfer of C from Betula papyrifera
to Pseudotsuga menziesii but the overall ecological
Plant Soil (2009) 321:5–33
significance of inter-plant C transfer has been questioned by Robinson and Fitter (1999). NMR studies
of common AM mycelial networks by Pfeffer et al.
(2004) revealed that, although significant amounts of
C were transferred between different roots connected
by a common fungal mycelium, the labelled C
remained within fungal compounds and no transfer
of C from fungus to plant took place. As pointed out
by Pfeffer et al. (2004) and earlier by Finlay and
Söderström (1992) such distribution of C within
mycelial networks may be of significance even in the
absence of net transfer of C from fungus to plants since
it would reduce the C demand of the fungal mycelium
colonising newly connected host plants and enable them
to gain access to nutrients taken up by the mycelium.
Although the predominant movement of C in fully
autotrophic mycorrhizal hosts is likely to be from plant
to fungus, over 400 plant species are achlorophyllous
and described as ‘myco-heterotrophic’, obtaining their
C from fungi. DNA-based studies of these fungi have
revealed most of them to be mycorrhizal species
colonising other autotrophic plants. The mycoheterotrophic species are thus effectively ‘cheaters’ or
epiparasites obtaining their C and nutrients through
mycorrhizal connections with neighbouring autotrophic plants (Bidartondo 2005; Bidartondo et al. 2002;
Leake 2004). In orchids the direction of C transfer is
often reversed since about 100 species are completely
achlorophyllous and all others pass through a germination and early developmental phase in which they
are dependent on an external supply of nutrients and C
since they have minute, dust-like seeds with no
reserves. Survival of germinating seedlings is thus
dependent upon rapid integration into fungal mycelial
networks. Although this pathway of C transfer is
sometimes dismissed as a ‘special case’ in discussions
concerning the overall significance of C transfer via
mycorrhizal hyphal connections, the Orchidaceae is the
largest family in the plant kingdom with over 30,000
species so the habit is arguably widespread and of
evolutionary significance.
Acquisition of N (Bending and Read 1995) and P
(Lindahl et al. 2001) by ectomycorrhizal fungi
colonising organic substrates is dependent on resources allocated to the mycelium. Ectomycorrhizal and
ericoid mycorrhizal fungi play a pivotal role in the
mobilisation of N and P from organic polymers (Read
and Perez-Moreno 2003) and their enzymatic capacities
have been reviewed by Lindahl et al. (2005).
13
Increased ectomycorrhizal mycelial growth and biomass production, resulting in selective spatial allocation of C to nutrient rich substrates has been
demonstrated in a range of studies (see Read and
Perez-Moreno 2003) and been shown to be associated with mobilisation of N and P. Energy is undoubtedly required for the synthesis of enzymes involved
in the mobilisation of nutrients but the partitioning of
C between fungal biomass production and hydrolytic
activity is not yet fully understood. Experiments by
Lindahl et al. (2007) suggest that decomposition of
litter by saprotrophs and mobilisation of N from welldecomposed organic matter may be spatially and
temporally separated in boreal forests. Many of the
organic N compounds taken up by ectomycorrhizal
mycelium contain C derived from photosynthetic
products originally translocated to the soil via the
same mycelium. This may reduce the C drain
imposed upon the host plant by ectomycorrhizal
symbionts. In axenically grown Betula pendula
plants supplied with 14C labelled protein as the sole
exogenous N source, only ectomycorrhizal plants
were able to exploit this N source. Heterotrophic
uptake of C associated with utilisation of this organic
N source was estimated to be up to 9% of plant C
over a 55 day period (Abuzinadah and Read 1989).
Simple amino acid sources are taken up intact by a
range of mycorrhizal plants as demonstrated in field
experiments by (Näsholm et al. 1998) and this also
contributes to the reverse flow of C through the
rhizosphere to plant roots. Utilisation of organic N
sources by arbuscular mycorrhizal plants is less well
understood but Hodge et al. (2001) demonstrated
enhanced decomposition and capture of N from
decaying grass leaves in the presence of AM fungi.
Further experiments are needed to distinguish between direct capture and uptake of organic N by the
hyphae and indirect uptake of inorganic N through
enhanced decomposition. It is possible that mycorrhizal hyphae contribute to rhizosphere priming via a
release of energy rich C which is utilised by
microbial saprotrophs. The mycorrhizal mycelium
provides a vastly increased surface area (compared
with roots alone) for interactions with other microorganisms and an important pathway for translocation into the soil of energy-rich compounds derived
from plant assimilates. Soluble C compounds released by the extraradical mycelium of arbuscular
fungi have been shown to influence the activity of
14
both fungi and bacteria associated with the mycorrhizosphere (Filion et al. 1999; Toljander et al. 2007).
Both stimulatory and inhibitory interactions are
possible and these have been reviewed with respect
to their relevance in sustainable agriculture by
Johansson et al. (2004). Production of mycorrhizal
mycelial exudates has been shown to influence
bacterial species composition and vitality (Toljander
et al. 2007) and vitality of mycorrhizal hyphae in turn
has been shown to influence attachment of different
bacteria to AM hyphae (Toljander et al. 2006). Other
recent experiments indicate that AM fungi may
influence bacterial assemblages in roots but that the
effect is not reciprocal (Singh et al. 2008). AM fungi
also produce a glycoprotein, glomalin, which is
deposited in soil as hyphae senesce and has been
estimated to constitute as much as 5% of soil C (see
Treseder and Turner 2007). As well as playing a role
in soil aggregation glomalin production is thought to
sequester significant amounts of C on a global scale
(Treseder and Turner 2007).
Exudation and reabsorption of some C compounds
from fluid droplets produced at ectomycorrhizal
hyphal tips has been demonstrated by Sun et al.
(1999) who concluded that it might represent an
important mechanism for conditioning the hyphal
environment in the vicinity of tips, creating an
interface for the exchange of nutrients and C
compounds with the adjacent soil environment and
its other micro-organisms. Ectomycorrhizal fungi
produce significant amounts of organic acids (Sun et
al. 1999; Ahonen-Jonnarth et al. 2000) which may
play a role in weathering of minerals, complexation
of toxic Al 3+ or in antibiosis. The microbial
decomposition of these organic acids could also
contribute significantly to soil respiration (van Hees
et al. 2005). Experiments by Rosling et al. (2004a, b)
suggest that mycorrhizal and other fungi differ in
their ability to allocate C to different mineral substrates
and that more labelled C is allocated to easily weatherable minerals such as potassium feldspar than to
quartz.
Despite the fact that the rhizosphere is defined in
terms of its elevated levels of soil microbiological
activity, we still know surprisingly little about the role
of rhizosphere communities in C flow, and little is
known about the roles of different members of the
community in assimilating plant exudates. Experiments by Ostle et al. (2003) and Rangel-Castro et al.
Plant Soil (2009) 321:5–33
(2005a) demonstrated rapid allocation and incorporation of recently photosynthesized 13C into soil
microbial biomass. Labelled C is incorporated within
hours and the half life of microbial pools of 13C was
calculated to be 4.7 days. RNA-based stable isotope
probing experiments by Rangel-Castro et al. (2005b)
using DGGE analysis of bacterial, fungal and archaea,
showed that active communities in limed soils were
more complex than those in unlimed soils and were
more active in utilization of recently exuded 13C
compounds. This suggests that in unlimed soils the
active microbial community may have been utilizing
other sources of C but the results may also reflect
differences in the amount of root exudation in limed
and unlimed grasslands. Another approach which has
been used to study bacterial communities associated
with mycorrhizal and non-mycorrhizal root systems is
the use of symbiosis-defective plant mutants. In
experiments by Offre et al. (2008), Oxalobacteraceae
isolates were more abundant in mycorrhizal roots of
Medicago truncatula than in non-mycorrhizal roots of
symbiosis-defective plants, whereas Comamonadaceae
isolates were more abundant in non-mycorrhizal roots.
New approaches based on stable isotope probing,
RNA analysis, and metagenomics (Vandenkoornhuyse
et al. 2007) indicate that there are many hitherto
unidentified root symbionts and that bacteria and AM
fungi occupying roots show differential activity in C
consumption with much higher C flow to some fungi
than others. Therefore, while it is clear that symbionts
are important determinants of rhizodeposition, our
understanding remains poor in many respects. While
this article is about C flow in the rhizosphere, and there
has been a general tendency in rhizosphere research to
concentrate on “quantitatively significant” C fluxes, it
should be remembered that plants produce a wide
spectrum of chemicals which are usually called
secondary metabolites because of their presumed
secondary role in plant growth. Chemicals released in
the rhizosphere play vital roles in signalling between
plant roots and different microorganisms. Although
these chemicals may only constitute a small proportion
of the total photosynthetically derived C flow from
roots they can play a key role in plant survival through
defence against pathogens or in attracting beneficial
symbionts. One example of this is the strigolactones,
that are produced in the root exudates of many
monocot and dicot species (Bouwmeester et al.
2007). These compounds induce branching of arbus-
Plant Soil (2009) 321:5–33
cular mycorrhizal fungi but also stimulate the germination of seeds of parasitic plants (Striga and Orobanche spp.). However, infection by Striga is reduced
in plants colonised by AM fungi through downregulating the production of the germination stimulant.
Phosphate starvation is known to induce strigolactone
production, and also to favour AM colonisation, while
AM fungi are known to improve the P status of their
hosts, which in turn would repress strigolactone
production. The effects of environmental factors on
numerous other signalling molecules are still entirely
unknown, although their effects on plant growth and
survival may be of paramount importance. Therefore,
although more quantitative studies of C and N flux
in the rhizosphere are still needed, these should also
be complemented by further qualitative studies of the
role of different signalling molecules, the roles these
play in plant–soil–microbe interactions and the way
in which they are influenced by different environmental conditions.
Carbon flow in the rhizosphere is bi-directional
Prior to 1990, the general consensus was that
rhizodeposition was a unidirectional flux whereby
plant C was lost from roots into the soil (Curl and
Truelove 1986). Once in the soil it was assumed to
undergo a number of fates including movement away
from the root in the soil solution due to diffusion and
mass flow, capture by soil microorganisms, and
sorption to the solid (Martin 1975; Newman and
Watson 1977). However, experiments undertaken in
hydroponic culture and subsequently soil revealed
that plant roots can also take up a range of organic
compounds from the soil into the roots with subsequent transfer to the shoots (Jones and Darrah 1992,
1993, 1994). Of the compounds investigated so far,
roots from a range of species have been shown to take
up predominantly low molecular weight solutes such
as organic acids, sugars and amino acids (Jones and
Darrah 1995; Sacchi et al. 2000; Thornton 2001). In
addition, roots may also take up inorganic C from
outside the root when present in a dissolved form (e.g.
HCO3−; Cram 1974; Amiro and Ewing 1992; Ford et
al. 2007). Although HCO3− can be readily converted
to organic acids inside the root, the contribution of
this inwardly directed inorganic C flux to the overall
C economy of the plants is small especially in view of
15
the large amount of HCO3− generated in respiratory
processes (Ford et al. 2007). One potential exception
occurs within proteoid roots of lupin roots where
significant uptake and assimilation of HCO3− into
malate and citrate occurs (Johnson et al. 1996). These
HCO3− derived organic acids are then exuded back
into the soil to aid in P mobilization in the rhizosphere.
Discrimination also needs to be made between
organic C that is taken up and assimilated in a
controlled (i.e. active transport) way and that which is
inadvertently taken up as a consequence of its
physicochemical properties (i.e. passive transport).
In the case of compounds with a high octanol–water
partition coefficient (KOW) value, these can simply
become sorbed to cell membranes and subsequently
metabolised (e.g. pesticides, chlorinated hydrocarbons; Scheunert et al. 1994). This passive process
can be expected to have no positive benefit to the
plant. Similarly, positively charged organic compounds can become sorbed to cell walls with no
subsequent assimilation. Some neutrally charged
compounds (e.g. acetic acid) can also passively
enter the cell if the concentration outside is greater
than that inside. While this has been used as an
experimental tool to understand membrane function
its significance in soil remains unknown (Herrmann
and Felle 1995).
Of greatest ecological significance is the active
root uptake of sugars and organic nitrogen compounds (e.g. amino acids, polyamines etc) from soil.
Typically, these compounds are taken into the plant
by co-transporters which are constitutively expressed
and located throughout the root system (Jones and
Darrah 1994, 1996; Fig. 3). These co-transporters are
powered by the plasma membrane H+-ATPases which
are predominantly located in the epidermis rather than
in the root cortex although levels of H+-ATPases are
also high in the stellar regions (Samuels et al. 1992;
Jahn et al. 1998). The transport proteins simultaneously transport H+ across the plasma membrane
together with individual organic solutes. The transporters are also relatively solute specific with transport families for amino acids and sugars being well
characterised at both the physiological and molecular
level (Fischer et al. 1998; Williams et al. 2000; Hirner
et al. 2006). In addition, membrane transporters also
exist for other solutes such as peptides, flavonoids
and polyamines although these protein families
remain less well characterised (DiTomaso et al.
16
Plant Soil (2009) 321:5–33
1992; Hart et al. 1992; Buer et al. 2007; Jones et al.
2005a, b). There is also strong evidence to suggest
that plant roots can take up larger molecular weight
solutes by endocytosis (Samaj et al. 2005). Current
evidence suggest that this process is important for
auxin-mediated cell–cell communication, polar
growth, gravitropic responses, cytokinesis and cell
wall morphogenesis (Ovecka et al. 2005).
As the plant expends energy in the uptake of these
compounds from soil we assume that the process
must confer some benefit to the plant. At present there
are four principal hypotheses to explain why plants
might take up organic solutes from soil (Fig. 4).
Although there is no reason to suggest that these are
mutually exclusive it is likely that their importance
varies in space and time within a root system and
between plant species.
Hypothesis 1: direct root exudate recapture
The first explanation is that the root is simply
recapturing C back from the soil that it previously
lost in response to passive root exudation, the latter
being a process over which it exerts little direct
control (Jones et al. 1996). This recapture of exudate
C not only enhances C use efficiency in the plant but
Internal C transport
1
4
2
Exudate
recapture
Signal
exchange
3
also prevents C accumulation in the rhizosphere
thereby reducing the growth of the soil microbial
community. This may serve three purposes by (1)
reducing microbial competition for poorly available
nutrients required by the root (e.g. N and P), (2)
reducing the growth of potentially pathogenic organisms, and (3) minimizing chemotactic gradients for
pathogenic organisms. When chemotaxis of beneficial
organisms is required, current evidence suggests that
more chemically specific signals at low concentrations are released in root exudates in a spatially and
temporally controlled manner (e.g. flavonoids;
Antunes et al. 2006; Sugiyama et al. 2007).
Hypothesis 2: indirect, fortuitous root exudate
recapture
The second explanation is that re-uptake of C from
soil might simply be indirectly related to normal
source-sink C delivery mechanisms in plants. In most
roots, solutes arriving from the shoots are unloaded
symplastically from the phloem, however, some
subsequently leak into the apoplast where retrieval
by active transporters can occur (Eleftheriou and
Lazarou 1997; Patrick 1997). This is unlikely to be
of significance in areas with a well developed
exodermis and tissues with high symplastic connectivity, however, it may be important in root caps and
cells where plasmodesmata have been blocked (Zhu
and Rost 2000; Hukin et al. 2002). These re-uptake
processes may also be indirectly linked to cell wall
bound invertases (Huang et al. 2007). These enzymes
convert apoplastic sucrose to glucose and fructose
which are then taken into the cell by co-localized
sugar transporters (Dimou et al. 2005). Import of
extracellular hexose sugars has been linked to a range
of sensing and signalling pathways in addition to their
potential role in supplying sugars for cellular expansion (Sherson et al. 2003).
Organic nutrient uptake
Hypothesis 3: nutrient capture from soil
Fig. 4 Schematic representation of a transverse root section
illustrating the four principal hypotheses explaining the uptake
of organic C from soil: 1 indirect, fortuitous root exudate
recapture in the root’s internal apoplastic transport and
signalling pathways, 2 direct recapture of root exudates from
the soil with the aim of reducing microbial growth and
pathogen chemotaxis, 3 uptake of organic nutrients (e.g. amino
acids) released during the mineralization of soil organic matter
in the rhizosphere, and 4 transfer of chemical signals involved
in inter-root and root–microbial communication pathways
The third explanation is that the uptake of organic
compounds from the soil may be a mechanism to
supply organic nutrients in addition to traditional
inorganic uptake routes (i.e. NO3−, NH4+, H2PO4−
etc). This may be particularly relevant in situations
where the supply of inorganic nutrients is limiting due
to either their low intrinsic solubility (e.g. P), low rate
Plant Soil (2009) 321:5–33
of ecosystem addition or a block in organic matter
mineralization preventing their release back into the
soil (e.g. N). It may also be particularly relevant to
non-mycorrhizal plants which lack the capability to
directly mineralize organic matter. Addition of isotopically labeled organic compounds to soil (15N, 13C,
14
C) has shown that roots have the potential to take up
and assimilate a wide range of compounds. In
agricultural soils, however, it has been shown that
plants are poor competitors for amino acids and
sugars in comparison to the soil microbial community
(Owen and Jones 2001; Bardgett et al. 2003;
Kuzyakov and Jones 2006). Consequently, unless
concentrations of organic solutes in the soil are very
high the uptake of exogenous organic N is likely to be
of minimal significance (Jones et al. 2005a, b). In
contrast, work in predominantly arctic and alpine soils
has suggested that organic N taken up from the soil in
the form of amino acids may contribute significantly
to a plant’s N budget (Chapin et al. 1993). In this case
the direct uptake of organic N circumnavigates the
need for the soil microbial community to mineralize
soil organic matter (Lipson and Nasholm 2001). The
uptake of organic N by roots is often viewed in the
literature as being unidirectional. The lack of consideration for an outward flux (i.e. root exudation)
therefore brings into question many of the rates of
flux reported in the literature. In most experiments,
rates of organic N uptake into roots are measured with
dual 15N-13C labeled compounds. As exudation of
organic N is derived predominantly from nonisotopically labeled organic N, due to the large
internal organic N reservoir relative to the amount
added, most isotopic tracer experiments will greatly
overestimate the rates of uptake. What we really
require are not measurements of gross rates of uptake,
but moreover net rates of uptake (i.e. influx minus
efflux; Philips et al. 2006). In the case of amino acids
and sugars, measurements in sterile hydroponic culture
have shown that the point of net zero uptake (i.e. influx
= efflux) occurs when the external concentration is
between 0.5 and 10 μM (Jones and Darrah 1994,
1996; Phillips et al. 2004). In most situations this is
extremely similar to the concentrations which exist
naturally in soil solution (Andersson and Berggren
2005; Jones et al. 2005a, b; Boddy et al. 2007)
suggesting that the contribution of organic N uptake
from soil may be less important than as a mechanism
for retaining the resources it already has (i.e. recapture
17
of exudates). The interpretation of isotopic flux
measurements is also complicated by the knowledge
that some organic N compounds can be firstly broken
down in the soil and the 15N released taken up as
15
NH4+ or 15NO3−. Measurements of the relative
enrichment of 15N and 13C in the roots can potentially
be used to discriminate between 15N taken up in an
intact form versus that previously mineralized in the
soil. However, after amino acids enter the root they can
undergo a number of metabolic reactions that can
ultimately lead to approximately 40–60% of the 13C
being released as 13CO2 (e.g. transamination and
deamination; Owen and Jones 2001). Similarly, the
loss of organic N derived CO2 can also occur after
uptake by mycorrhizas again leading to an underestimation of the organic N flux. Consequently, isotopic
flux measurements are fraught with potential pitfalls
that make interpretation of organic C and N fluxes at
the root–soil interface extremely difficult (Jones et al.
2005a, b).
Hypothesis 4: rhizosphere signalling
The fourth explanation for plants actively taking up
organic compounds from soil is for inter and intraroot signalling and for root–microbial signal exchange. In comparison to the other three potential
explanations, this is a poorly explored aspect of
rhizosphere ecology (Bais et al. 2004). Although
sugars are important in plant signalling, there is
currently no evidence to suggest that they would
provide specific signals to enable effective communication between roots and other organisms in the
rhizosphere. More likely is that root transporters
would be involved in the uptake of highly specific
signalling molecules (e.g. peptides). In other cases,
compounds released from microorganisms can have a
direct effect on plant growth and metabolism (Brown
1972), however, the mode of transport of these
signalling molecules into the root remains unknown
(e.g. lumichrome; Phillips et al. 2004; Matiru and
Dakora 2005). As our understanding of the diversity
and control of signalling processes in plants increases
it is likely that some of these will have functional
significance in the rhizosphere (Bais et al. 2004;
Bahyrycz and Konopinska 2007; Jun et al. 2008).
Further discussion of this issue can be found in
Hartmann et al. (2009), Lambers et al. (2009) and
Faure et al. (2009).
18
Rhizodeposition in the plant C and N budget
Methods of investigation
While rhizodeposition can be quantified relatively
easily in the absence of soil by growing roots in
sterile hydroponic culture and collecting the C
accumulating in the external media, this method lacks
ecological relevance (Ryan et al. 2001). Quantifying
rhizosphere C-flow in relation to soil environments,
however, has proved extremely difficult. The amount
of rhizodeposition entering soil during a growing
season typically represents only a small amount of C
and N in comparison to that already present in the soil
organic matter (SOM) and therefore measuring
changes in soil C in response to rhizodeposition
remains virtually impossible. This is also highly
pertinent to the uncertainties surrounding the effects
of environmental change on the soil C-balance where,
although significant effects on soil C-sequestration are
predicted, changes in soil C are difficult to detect due
to the large SOM-C background and high degree of
spatial variability in SOM. Consequently, tracing
root-derived C and N by isotopic techniques is a
prerequisite for the quantification of rhizodeposition
in soil (Warembourg and Kummerow 1991). For C, a
widely used technique involves the exposure of
shoots to a 13C or 14C enriched atmosphere to label
the photoassimilates. Subsequently, the fixed isotope
tracer becomes partitioned to range of operationally
defined below-ground compartments (roots, soil residues including microbial biomass, and root-derived
CO2). Changes in isotope abundance in these pools is
typically followed over time to estimate rhizodeposition as part of the plant or ecosystem C budget. The
experimental conditions have to be carefully considered when interpreting the partitioning of photoassimilates. For example, the length of the isotopic
labeling and subsequent chase period is a major
determinant of the amount of C delivered into the
soil (Meharg 1994). Short-term pulse labeling
(minutes to hours) traces rhizodeposits derived predominantly from recent photoassimilates (i.e. root
exudates, mucilage and border cells). Accordingly,
pulse-labeling by this method tends to underestimate
total rhizodeposition but remains useful in the
investigation of assimilate partitioning in relation to
plant metabolism (Phillips and Fahey 2005; Allard et
al. 2006; Hill et al. 2007). Longer isotopic labeling
Plant Soil (2009) 321:5–33
periods (weeks to months) trace not only the
senescence and turnover of roots but also the fraction
of root exudates that may not be derived from recent
C (Swinnen et al. 1995). In some cases this may be a
significant part of total root exudation (Thornton et al.
2004).
Most experiments in soil have tended to focus on
the total amount of C lost in rhizodeposition, while
hydroponic studies carried out in the laboratory have
tended to focus on the tracking of individual
compounds lost from roots. The isotopic tracking of
individual root-derived compounds into soil, however, has only recently become routinely possible
(Paterson et al. 2008). The use of gas chromatography
coupled to isotope ratio mass spectrometry allows the
dynamic tracking of specific rhizodeposits such as
sugars from roots and associated symbionts into soil
(Derrien et al. 2004; Paterson et al. 2007). Another
recent breakthrough for tracing C flow in the
rhizosphere is stable isotope probing (SIP). In this
technique, isotopically labeled plant assimilates are
released into the soil and then subsequently taken up
and incorporated into the soil microbial community.
The isotopically labeled microbial DNA, RNA or
phospholipids can then be extracted and their isotope
ratio determined whilst genetic material can be
sequenced to identify members of the soil microbial
community consuming the rhizodeposits (Singh et al.
2004, Rangel-Castro et al. 2005a, b; Shrestha et al.
2008). However, labeling of photoassimilates often
requires a sophisticated experimental set-up particularly for large plants (Warembourg and Kummerow
1991). In the case of 14C, its use may be problematic,
particularly in the field, due to safety and environmental concerns. This severely limits isotopic investigations on field-grown plants over their entire life
cycle (unless 13C pulse-labelling is used, Högberg et
al. 2008). Ultimately, this represents a serious concern
when calculating agro- or natural-ecosystem C budgets and the potential contribution of rhizodeposition to
C sequestration. The use of natural abundance of the
stable isotope 13C can provide an elegant alternative
(i.e. δ13C; Ekblad and Hogberg 2001). Growing a C4
plant on a C3 history soil (and vice versa) allows the
tracing of new plant-derived C from the C4-plant in
soil because of the difference in 13C natural abundance in plant material between C3 and C4 vegetation
(Boutton 1996; Rochette et al. 1999). However, this
approach is restricted to limited contexts, typically to
Plant Soil (2009) 321:5–33
maize grown on a C3 soil because of the difficulty to
find a soil with a known C3 or C4 history (Balesdent
and Balabane 1992; Qian et al. 1997). Non-isotopic
approaches for quantifying rhizodeposition are available using a range of microbial biosensors. These
have been employed as semi-quantitative measures of
total root C flow and more recently for spatially
localising the release of specific exudate components
(Paterson et al. 2006).
One major difficulty when attempting to quantify
the transfer of labeled C below ground is the
mineralization of rhizodeposits by rhizosphere microorganisms. Typically, low molecular weight (MW)
root exudates are believed to only have a residence
time of a few hours in soil solution as they are rapidly
consumed by the C-limited rhizosphere microbial
community (Nguyen and Guckert 2001; van Hees et
al. 2005). However, as it was observed for glucose,
the microbial uptake of substrate and its subsequent
mineralization may be decoupled in time. Therefore,
the turnover of low molecular weight exudates in soil
solution as determined from the kinetics of mineralization are likely to be underestimated by an order of
magnitude, indicating turnover times of minutes
rather than hours (Hill et al. 2008). Although higher
MW rhizodeposits have a slightly longer persistence
time in soil, they are still mineralized within a few
days (Mary et al. 1992, 1993; Nguyen et al. 2008).
This rapid biodegradation of rhizodeposits means that
a significant proportion of the rhizodeposits are quickly
lost from the soil as labeled CO2 (rhizomicrobial
respiration). The longer the labeling and the chase
period, the greater the amounts of rhizodeposits are
lost in this way. Because a large proportion of the
rhizodeposits are low MW and labile, rhizomicrobial
respiration overlaps directly in time and space with
root/symbiont respiration of the same labeled photoassimilates (Dilkes et al. 2004). Experimentally, the
flux of labeled CO2 derived directly from roots and
indirectly from rhizodeposits are therefore determined
together (rhizosphere respiration; Todorovic et al.
2001). Knowledge of the partitioning of rhizosphere
respiration into root, symbiont and rhizomicrobial
components, however, is crucial if we are to gain a
deeper understanding of rhizosphere C flow (Paterson
2003; Paterson et al. 2005). Many attempts have been
made to partition rhizosphere respiration, from the
simple use of antibiotics to more sophisticated models
based on isotopic methods, however, none has proved
19
satisfactory from a quantitative perspective (Cheng et
al. 1993; Kuzyakov 2006; Sapronov and Kuzyakov
2007). Current estimates suggest that approximately
50% of rhizosphere respiration is due to the turnover of
rhizodeposits and 50% to direct root (and mycorrhizal)
respiration, however, it is clear that this needs to be
major focus for research in the future (Kuzyakov
2002).
Estimating N rhizodeposition in soil is commonly
undertaken with a 15N isotopic tracer (Hertenberger
and Wanek 2004). The tracer is supplied by a foliar
application as a solution or spray, by stem feeding, by
a pre-culture on a labeled substrate or by using splitroot systems, one compartment for being used for the
labeling and the other for determining the release of
15
N from roots (Jensen 1996; Hogh-Jensen and
Schjoerring 2001; Mayer et al. 2003). All methods
currently assume, (1) a homogenous mixing of the
tracer within the plant N pool, and (2) that the
isotopic signature of N-rhizodeposits is the same as
that of the roots. Further rigorous validation of these
assumptions is required. Depending on the soil
conditions, a fraction of the rhizodeposited 15N also
may be unrecovered due to denitrification leading to
an underestimation of N rhizodeposition (de Graaff et
al. 2007). Recent advances have also been made in
the dual 13C-15N isotopic labeling of plants in situ
(Wichern et al. 2007).
To overcome the difficulties related to non-sterile
soil conditions, many studies were and are still
conducted in sterile hydroponic culture. Under these
conditions both the amount and the nature of
compounds released from roots can be determined.
However, one has to be aware of the limitations of
such experimental conditions. For instance, both the
nature and quantity of compounds released from roots
depends on the plant/root physiology, which greatly
differs between a simple sterile nutrient solution and a
complex soil environment (Neumann and Römheld
2001). Furthermore, exudation has been quantified for
decades in nutrient solution which are not regularly
renewed, a system that exacerbates the re-uptake of
exudates by roots and that leads to a large underestimation in exudation rates (Jones and Darrah 1993).
Consequently, it is necessary to adapt the experimental set-up used to study exudation so that it account
for the re-uptake of exudates. This can be done by
using microcosms percolated by nutrient solution
(Hodge et al. 1996) or by modelling the kinetics of
20
exudate accumulation in the root bathing solution as
the net output between the gross efflux and the reuptake of exudates (Personeni et al. 2007). The use of
bioreporter microorganisms is also an interesting
approach to spatially localize the release of some
specific compounds or class of compounds, however,
quantitative information about the rhizodeposition
flux are difficult to achieve (Yeomans et al. 1999;
Darwent et al. 2003).
Investigations of rhizodeposition are hampered by
many technical difficulties and sometimes by unresolved methodological problems arising from the
numerous interactions between roots, the soil matrix
and microorganisms (van Hees et al. 2005). Availability of robust methodologies for the qualitative and
quantitative determination of rhizodeposition in soil
clearly remains an unsolved issue. Current methods
are often incomplete or biased and consequently,
estimates of the flux of C (and to a lesser extend of N)
to the rhizosphere are associated with significant
uncertainty.
How much C is lost via rhizodeposition?
In the last few decades, hundreds of attempts have
been made to quantify the amount of photoassimilate
C partitioned below ground (Nguyen 2003). Most of
the initial studies used 14C although 13C is now
increasingly being used for tracing purposes. Results
are commonly expressed as partition coefficients
describing that amount of net fixed C allocated
between shoots, roots, rhizosphere respiration (root
and symbiont respiration + respiration of rhizodeposits) and soil residues. Soil residues include rhizodeposits, microbial biomass-C and metabolites derived
from rhizodeposits (including mycorrhizal hyphae)
but also fine roots debris that cannot be effectively
separated from the soil (e.g. root hairs, epidermal cells
etc). Figure 5 summarizes a review of whole plant C
partitioning averaged across a wide range of published studies and updates previous reviews (Bidel et
al. 2000; Nguyen 2003). Overall, it is clear that most
isotopic labeling studies have focussed on young
plants at a vegetative stage (typically <1 month old).
The focus on young plants is due to methodological
difficulties in growing and labeling plants to maturity
in controlled conditions. However, plant age has a
strong effect of the partitioning of photoassimilate to
the rhizosphere. For example, in annual plants pulse-
Plant Soil (2009) 321:5–33
labelled with 14C, when comparing two plant ages
ranging within the range 28–600 days of culture, the
partitioning of C to rhizosphere clearly decreases with
plant age of −43%, −28%, −20% for roots, rhizosphere respiration and soil residues, respectively
(median values; Nguyen 2003). Although C partitioning has been investigated in a wide range of plant
species, almost half of the published data are for
wheat and rye-grass and 76% of the studies are related
to five crop/grassland species. Hence, we currently
have a very incomplete picture of C rhizodeposition
particularly in mixed plant communities. In particular,
dicotyledonous plants have received little attention
and the amount of rhizodeposition by trees in natural
ecosystems remains virtually unknown with the
exception of recent studies by (Högberg et al. 2008).
Furthermore, when very young trees have been used
to study rhizodeposition, the experimental conditions
employed often bear little relevance to natural forest
stands. Consequently, our poor knowledge of rhizodeposition in trees is problematic, particularly when
quantifying C sequestration in forests.
Studies indicate that roughly 40% of net fixed C is
allocated belowground. For cereals and grasses, this
approximates to around 1.5–2.2 t C ha−1 for the
vegetation period (Kuzyakov and Domanski 2000).
Of the C partitioned below ground about 50% of it is
retained in root biomass (19% of net fixed C), 33% is
returned to the atmosphere as rhizosphere respiration
(12% of net fixed C), 12% can be recovered as soil
residues (5% of net fixed C) and a small amount is
lost by leaching and surface runoff. Assuming that
roots and microorganisms contribute equally to
rhizosphere respiration (Kuzyakov 2006), an assumption that must be treated with caution, then a rough
estimate of rhizodeposition would be around 11% of
the net fixed C or 27% of C allocated to roots. This
would correspond to 400–600 kg C ha−1 for the
vegetation period of grasses and cereals. These values
only provide a rough estimate, however, due to the
uncertainty surrounding the partitioning of rhizosphere respiration and because soil residues often
include small roots and living mycorrhizal mycelium
that cannot be realistically separated from soil by
current protocols. This probably explains the skewed
distribution of the soil residue partitioning coefficients
(Fig. 5).
Studies quantifying the amount of N rhizodeposition are much less numerous and a survey of the
Plant Soil (2009) 321:5–33
% of total
Shoots
27.7
20.3
10.3
9.6
8.5
3.3
3.0
3.0
1.8
1.8
1.5
1.5
1.5
1.1
1.1
0.7
0.7
0.7
0.7
0.7
0.4
100
100
40
60
% of total recovered
40
20
80
Belowground
100
80
60
40
20
0
Median=60%
Roots
% of tracer recovered
20
60
Days
100
80
60
40
20
0
Median=12%
Soilil residues
100
Median=33 days
Rhizosphere CO2
100
80
0
Plant age
0
CO 2
% of tracer recovered
Triticum aestivum
Lolium perenne
Bromus erectus
Hordeum vulgare
Zea mays
Avena sativa
Bromus madritensis
Castanea sativa
Salix viminalis
Trifolium repens
Festuca arundinacea
Pinus taeda
Populus tremuloides
Brassica napus
Festuca pratensis
Cynodon dactylon
Lolium multiflorum
Lycopersicon esculent
Medicago truncatula
Pisum sativum
Bouteloua gracilis
Total
Number of
partitioning
coefficient sets
75
55
28
26
23
9
8
8
5
5
4
4
4
3
3
2
2
2
2
2
1
271
% of tracer recovered
Species
21
Median=19%
80
60
40
20
0
Median=5%
Fig. 5 Partitioning of labeled net fixed C after a pulse or
continuous exposure of shoots to a 14CO2 enriched atmosphere.
For each compartment, boxplots show the distribution of 271
individual partition coefficients drawn from a review of the
literature by Nguyen (2003) and updated to 2007. The plant
species and the distribution of plant ages are provided on the
left. The box represents the second and third quartiles separated
by the median. The whiskers extend to 1.5 times the
interquartile range. The circles denote outliers
literature shows that our knowledge of this phenomenon is very incomplete (Fig. 6). More than 60% of
the available data pertain to pea and wheat. In these
studies, N rhizodeposition accounts for 10–16% of
total plant N with losses higher in legumes in
comparison to non-legume species. This observation
may be biased, however, as the legume based studies
have tended to use older plants (Fig. 6) where a larger
part of rhizodeposited N may be attributed to root
turnover. As free amino acids and proteins represent
only a minor component of root exudates (typically
1–2% of exudate-C; Kraffczyk et al. 1984; Jones and
Darrah 1993) we assume that they contribute little to
plant N rhizodeposition. We conclude that N rhizodeposition must be largely due to root turnover or
possibly to an efflux of labeled ammonium and/or
nitrate (Feng et al. 1994; Scheurwater et al. 1999).
Small root debris that cannot be separated by common
sampling protocols may also lead to an overestimation of N rhizodeposition.
Published reports also show that the partition
coefficients of C and N both below ground and to
rhizodeposition are highly variable. This illustrates
that plant species, plant ecotype/cultivar, age and
environmental conditions all exert a strong impact on
rhizodeposition. From conventional tracer experiments it is often difficult to conclude about how
rhizodeposition is affected by environmental conditions as the partitioning of rhizosphere respiration
between the root and the microbial components may
also be altered. However, when the partitioning of C
to root, to rhizosphere respiration and to soil residues
changes in the same way, some conclusions may be
drawn. Hence, it can be assumed that the percentage
22
% of
total
5
1.3
Legumes (n=42)
Non Legume (n=33)
3
Median=16.5%
Median=10%
60
80
Median=2
% of plant N
1.3
36.0
Median=4
2
1.3
Plant age rank
1.3
1.3
1.3
1.3
4
2.7
N rhizodeposition
40
1.3
100
4.0
1.3
1.3
20
1.3
1
2.7
1.3
26.7
8.0
0
Avena sativa
Cajanus cajan
Cicer arietinum
Glycine max
Hordeum vulgare
Lathyrus sativus
Lolium perenne
Lupinus albus
Lupinus angustifolius
Medicago sativa
Ornithopus compressus
Pisum sativum
Trifolium pratense
Trifolium pratense
Trifolium repens
Trifolium subterraneum
Triticum aestivum
Triticum turgidum
Vicia faba
Vigna radiata
Total
Number of
results
3
1
1
1
2
1
1
1
1
1
1
27
1
1
2
1
20
6
2
1
75
0
Species
Plant Soil (2009) 321:5–33
Legumes (n=42)
Non Legume (n=33)
2.7
1.3
100
Fig. 6 Summary of published studies on N rhizodeposition
expressed as a percentage of total plant N. Rhizosphere N
derived from roots was determined by labeling of plant N with
15
N supplied as 15NH3, 15NO3 or 15N-urea. The technique used
was one of the following: split-root cultures, stem/petiole
infiltration/injection, leaf dipping, 15NH3-enriched atmosphere
or preculture on a 15N-labelled substrate. The plant species and
the ranked distribution of plant age are given on the left. Ranks
for plant ages are defined as follows: 1 early vegetative stage, 2
end of vegetative stage, 3 flowering/grain filling, 4 maturity.
The box represents the second and third quartiles separated by
the median. The whiskers extend to 1.5 times the interquartile
range. Circles represent outliers
of assimilates ending up as rhizodeposition generally
decreases with plant age and is increased by the
presence of microorganisms and by elevated atmospheric CO2.
We now have almost 30 years of knowledge from
C rhizodeposition research. From tracer experiments,
we can reasonably predict the order of magnitude of
this C flux for agroecosystems. These studies all attest
to rhizodeposition being a major C flux. In hindsight,
however, it is also evident that a quantitative approach
to assessing the functional role of rhizodeposition in
soil is strongly limited by technical difficulties arising
from the complex interactions occurring in the
rhizosphere and the tight link between rhizodeposition
and the plant’s physiological status. Accordingly,
there is an urgent need to develop new approaches
and methods for probing rhizodeposition. The coupling of plant labeling with molecular tools is
promising for understanding the link between the
plant-derived C and microbial processes in the
rhizosphere but the current information remains more
qualitative than quantitative. Considering the need to
have a quantitative understanding of C and N fluxes
in the rhizosphere to predict ecosystem behaviour,
modelling approaches should be considered to be of
major importance. For example, integrated modelling
of rhizosphere functioning could help to assess
previous estimates of rhizodeposition by cross validation of rhizodeposition models with other models,
for which the output variables are tightly connected to
rhizodeposition and are more accessible (e.g. microbial growth, N dynamics). This could help to integrate
our knowledge, to link rhizodeposition with plant
functioning and to upscale case studies to the
ecosystem level.
Modelling approaches
Mathematical modelling has the potential to predict C
flows at spatial and temporal scales that are beyond
the capability of current experimental techniques
(Darrah et al. 2006). The construction and use of
these models, however, are only as good as the
knowledge of the individual processes and the values
they are parameterized with. We know that the
rhizosphere is inherently complex and that by default,
current mathematical models are highly simplistic
from a mechanistic standpoint. Despite this, however,
there is also no doubt that they have greatly improved
our understanding of rhizosphere processes (Barber
1995; Nye and Tinker 2000). In addition, it is also
clear that major advances in mathematically describ-
Plant Soil (2009) 321:5–33
23
ing the complexity of the rhizosphere have been made
in recent years (Roose and Fowler 2004; Schnepf and
Roose 2006). These advances have been only become
possible through interdisciplinary interaction between
applied mathematicians and rhizosphere biologists.
In a rhizodeposition context, one of the first
quantitative modelling approaches was that taken by
Newman and Watson (1977) where rhizosphere C flow
was used to drive a soil microbial growth model. This
model was subsequently refined by Darrah (1991a, b)
with microbial growth placed in a growing root
context. In terms of whole plant modelling, a
photosynthesis model was used to calculate the flux
of C entering the soil using photoassimilate partition
coefficients (Swinnen 1994). However, due to the tight
relationship between rhizodeposition and plant physiology, the input of C into the soil is not a constant
part of the net fixed C or even of the C allocated to
roots (see above). Therefore, it is necessary to have a
more mechanistical approach, by modelling rhizodeposition along with plant physiology and more
particularly with root system functioning. Indeed,
exudation, which is a major component of rhizodeposition, is dependent upon root surface area and on the
C concentration in root tissue relative to that in the soil
solution. Subsequently, exudation can be simplistically
modelled by a diffusion equation placed in a vegetation model that simulates plant phenology, canopy
assimilation and carbohydrate partitioning above and
belowground (Grant 1993). Due to the higher rates of
exudation at root apices (McCully and Canny 1985;
Darwent et al. 2003), the number and type of lateral
branching is an important characteristic to be considered (Henry et al. 2005). Figure 7 shows an example
of how this can be done. Exudation of an individual
root was modelled from the root surface area (given by
the root length and diameter) and by including the
longitudinal variability of the C efflux (Personeni et al.
2007). Upscaling this model to the whole root system
was achieved by coupling the exudation model to a
root architecture model that simulates root emergence,
their length and diameter as a function of thermal time
(Pages and Pellerin 1996). When this was done the
simulated cumulative exudation was 4.9 g C plant−1 or
390 kg C ha−1 (eight plant m−2) at 860 growing degree
days (flowering). This estimate accounts for the
longitudinal variability of C efflux along individual
roots, for the number of branches and for the root
surface area of a model maize root system. This value
is consistent with rhizodeposition estimates from tracer
experiments, provided that rhizodeposition includes
not only exudation but also mucilage, border cells and
lysed cells.
There is great interest in coupling the modelling of
rhizodeposition with root architecture models as it
allows users to simulate changes in rhizodeposition in
response to environmental conditions or photoassimilate availability through modifications of the characteristics of the root system. For example, N availability
commonly increases root branching and consequently
the number of root apices with higher rates of
exudation. Similarly, limitation in C allocation to roots
induces a reduction in root branching and in root
diameter (Thaler and Pages 1998; Bidel et al. 2000)
Whole root system
Cumulative Exudation (g C plant -1)
4
h
4.9 g C plant-1
5 .2
Φ E ( x) =
(1 + x )0.43
5
-2 -1)
6
Single root efflux of C
Whole roots
390 kg C ha-1 (8 pl m-2)
10% net fixed C
3
290 mg C g-1 root DW
Secondary roots
2
Tertiary roots
0
1
Primary roots
0
5
10
15
20
Distance to the apex (cm)
25
Fig. 7 Modelling of root exudation in maize. Left: Experimentally parameterized efflux profile of C from a single root (from
Personeni et al. 2007). Right: Mathematical simulation of whole
root system exudation in maize from germination until flower-
Growing degree-days
ing (860 growing degree days, base 6°C). The simulation was
performed by coupling the single root efflux of C model to the
root architecture model of Pages and Pellerin (1996)
24
and consequently exudation. Furthermore, recent root
architecture models have also included C availability
in root tissue (Thaler and Pages 1998; Bidel et al.
2000), which is potentially important for modelling
diffusive losses. Therefore, further investigations are
needed to elucidate if a change in rhizodeposition
occurring in response to a modification in photoassimilate availability (Dilkes et al. 2004) is related to
changes in root architecture and/or to changes in C
availability within root tissues, which would change
the rhizodeposition by individual roots. Much work
has yet to be done to understand the mechanisms
of N release from roots but a similar approach to
that presented for C can be considered to model
rhizodeposition of N in relation to the root system
structure and functioning. Hence, modelling rhizodeposition with root architecture models that
integrate C and N availability in root tissue is
undoubtedly a promising perspective for predicting
the release of C and N by roots under various
environmental conditions.
Rhizodeposition—future outlook
It is clear from the previous discussion that we have
made great progress in highlighting the importance of
rhizosphere C flow in numerous aspects of ecosystem
functioning. However, it is also apparent that we have
a very long way to go before we can realistically
harness the full extent of this knowledge for landscape level management (e.g. forest sustainability,
biodiversity enhancement etc). This is exemplified by
our process level understanding of C and nutrient
flow at the single root level, however, how this scales
up in landscapes which contain a mosaic of hydrologically interconnected vegetation and soil types
remains unknown. This is certainly a goal which will
only occur through an integration and enhancement of
mathematical model scaling techniques. Indeed, the
rhizosphere can be expected to play a major role in all
the major challenges facing the planet including
greenhouse gas mitigation, sustainable food production
and food security, bioenergy production, preservation of
water quality, accelerated restoration of post-industrial
sites etc. One of the major obstacles to achieving this is
the shear complexity of the rhizosphere and the lack of
experimental techniques for teasing apart the myriad of
interactions between roots and their biological, chemical
Plant Soil (2009) 321:5–33
and physical environment. While our knowledge of
rhizodeposition has focused on crop plants, for both
practical and economic reasons, there is a critical need
to assess rhizosphere C flow in complex plant communities. A broader understanding of rhizosphere
responses throughout the plant world can yield great
insights into plant–soil functioning which cannot be
provided by working on crop plants alone (Lambers et
al. 2008). Similarly, there is also a need to look at
rhizosphere processes in mature trees and particularly
in mixed plantations where many synergistic relationships have been reported to occur (e.g. in litter
decomposition, mycorrhizal interactions etc; Rothe
and Binkley 2001). With the ongoing advances in our
experimental and theoretical understanding of plant
and microbial genomics, proteomics and metabolomics
and the current focus on systems biology (Meldrum
2000), it is evident that rhizodeposition will remain a
major focus of research for the foreseeable future. This
increase in technology inevitably brings new challenges.
In particular, finding robust statistical approaches to
disentangle massive datasets produced from temporal
and spatial sampling will be a necessity to maximise the
potential of the technology (e.g. those produced by
biodiversity measures such as pyrosequencing; Emerson
et al. 2008; Fulthorpe et al. 2008). Consequently,
rhizosphere bioinformatics is likely to grow in
importance in the next decade. One of the most
promising areas for further development is manipulating rhizosphere C flow to produce sustainable
agricultural production systems. If we can interpret
the C signals in the rhizosphere and then manipulate
their flow, there is a potential to influence rhizosphere
development. This could help to reduce our reliance
on pesticides if we can stimulate and preserve the
activity of biocontrol agents in the rhizosphere.
However, a note of caution must also be made in
our attempts to manipulate rhizosphere C flow.
Although there are many researchers around the
world attempting to alter rhizodeposition to help
reduce our over-reliance on chemical fertilizers and
pesticides, we must be careful not to over-mine or
exploit the natural resources to a point at which the
soils are left in a highly degraded state (i.e. unsuitable
for colonization by native plants by excessively
stripping the soil P pool). Consequently, there is also
a pressing need for a debate on the ethics of
manipulating the rhizosphere if we want to preserve
public support for our research.
Plant Soil (2009) 321:5–33
Acknowledgements The authors would like to address special
thanks to L. Pagès (INRA, Avignon) for providing simulations
from root architecture models.
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